Accepted Manuscript Isolation of two insecticidal toxins from venom of the Australian theraphosid spider Coremiocnemis tropix Maria P. Ikonomopoulou, Jennifer J. Smith, Volker Herzig, Sandy S. Pineda, Sławomir Dziemborowicz, Sing-Yan Er, Thomas Durek, John Gilchrist, Paul F. Alewood, Graham M. Nicholson, Frank Bosmans, Glenn F. King PII: S0041-0101(16)30314-2 DOI: 10.1016/j.toxicon.2016.10.013 Reference: TOXCON 5484 To appear in: Toxicon Received Date: 1 August 2016 Revised Date: 21 October 2016 Accepted Date: 25 October 2016 Please cite this article as: Ikonomopoulou, M.P., Smith, J.J., Herzig, V., Pineda, S.S., Dziemborowicz, S., Er, S.-Y., Durek, T., Gilchrist, J., Alewood, P.F., Nicholson, G.M., Bosmans, F., King, G.F., Isolation of two insecticidal toxins from venom of the Australian theraphosid spider Coremiocnemis tropix, Toxicon (2016), doi: 10.1016/j.toxicon.2016.10.013. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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Accepted Manuscript
Isolation of two insecticidal toxins from venom of the Australian theraphosid spiderCoremiocnemis tropix
Maria P. Ikonomopoulou, Jennifer J. Smith, Volker Herzig, Sandy S. Pineda,Sławomir Dziemborowicz, Sing-Yan Er, Thomas Durek, John Gilchrist, Paul F.Alewood, Graham M. Nicholson, Frank Bosmans, Glenn F. King
PII: S0041-0101(16)30314-2
DOI: 10.1016/j.toxicon.2016.10.013
Reference: TOXCON 5484
To appear in: Toxicon
Received Date: 1 August 2016
Revised Date: 21 October 2016
Accepted Date: 25 October 2016
Please cite this article as: Ikonomopoulou, M.P., Smith, J.J., Herzig, V., Pineda, S.S., Dziemborowicz,S., Er, S.-Y., Durek, T., Gilchrist, J., Alewood, P.F., Nicholson, G.M., Bosmans, F., King, G.F., Isolationof two insecticidal toxins from venom of the Australian theraphosid spider Coremiocnemis tropix, Toxicon(2016), doi: 10.1016/j.toxicon.2016.10.013.
This is a PDF file of an unedited manuscript that has been accepted for publication. As a service toour customers we are providing this early version of the manuscript. The manuscript will undergocopyediting, typesetting, and review of the resulting proof before it is published in its final form. Pleasenote that during the production process errors may be discovered which could affect the content, and alllegal disclaimers that apply to the journal pertain.
Isolation of two insecticidal toxins from venom of the
Australian theraphosid spider Coremiocnemis tropix
Maria P. Ikonomopoulou*§1, Jennifer J. Smith*1, Volker Herzig*1, Sandy S. Pineda1,
Sławomir Dziemborowicz2, Sing-Yan Er1, Thomas Durek1, John Gilchrist3, Paul F. Alewood1,
Graham M. Nicholson2, Frank Bosmans3, and Glenn F. King#1
1Institute for Molecular Bioscience, The University of Queensland, St. Lucia, QLD 4072,
Australia;
2School of Life Sciences, University of Technology Sydney, NSW 2007, Australia; 3Department of Physiology & Solomon H. Snyder Department of Neuroscience, Johns
Hopkins University, School of Medicine, Baltimore MD 21205, USA
* equal contributions
§ Current address: QIMR Berghofer Medical Research Institute, Herston QLD 4006, Australia
#Address correspondence: Prof. Glenn F. King, Institute for Molecular Bioscience, The
University of Queensland, 306 Carmody Road, St. Lucia, QLD 4072, Australia; Phone: +61 7
(EGTA) 5, and ATP-Na2 3, adjusted to pH 7.4 with 1 M CsOH. To eliminate any influence of
differences in osmotic pressure, all internal and external solutions were adjusted to 400 ± 5
mOsmol/L with sucrose. Experiments were rejected if leak currents exceeded 1 nA or currents
showed signs of poor space clamping.
Peak current amplitude was analysed offline using AxoGraph X v1.5.3 (Molecular Devices).
All curve-fitting was performed using PRISM 6 for Windows (GraphPad Software Inc., San
Diego, CA). Nonlinear least-squares regression was used to fit I/V curves whilst linear least-
squares regression was used to fit voltage-dependent block data. Comparisons of two sample
means were made using a paired Student’s t-test. A test was considered to be significant when
p < 0.05. All data represent the mean ± SEM of n independent experiments.
2.12 Activity of Ct1a on cloned Blattella germanica sodium channels.
The DNA sequence of the B. germanica NaV channel (BgNaV1) (Dong, 1997) was confirmed
by automated DNA sequencing and cRNA was synthesized using T7 polymerase (mMessage
mMachine kit, Life Technologies) after linearizing DNA with NotI. BgNaV1 was expressed in
Xenopus oocytes together with the TipE subunit (Feng et al., 1995) (1:5 molar ratio). After
cRNA injection, oocytes were incubated at 17°C (in 96 mM NaCl, 2 mM KCl, 5 mM HEPES,
1 mM MgCl2 and 1.8 mM CaCl2, 50 µg/mL gentamycin, pH 7.6 with NaOH) for 1–2 days,
then currents were measured using two-electrode voltage-clamp recording techniques (OC-
725C, Warner Instruments, Hamden, CT, USA) with a 150-µL recording chamber. Data were
filtered at 4 kHz and digitized at 20 kHz using pCLAMP software (Molecular Devices).
Microelectrode resistances were 0.5–1 MΩ when filled with 3M KCl. The external recording
solution contained (in mM): 96 NaCl, 2 KCl, 5 HEPES, 1 MgCl2 and 1.8 CaCl2, pH 7.6 with
NaOH. All experiments were performed at room temperature (~22°C). Leak and background
conductances, identified by blocking the channel with tetrodotoxin (TTX), were subtracted
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for currents shown.
Voltage–activation relationships were obtained by measuring steady-state currents and
calculating conductance. In brief, oocytes were depolarized in 5 mV increments from –90 mV
to 5 mV for 50 ms from a holding potential of –90 mV, followed by a depolarizing pulse to
–15 mV for 50 ms. Peak current from these steps was converted to conductance and
normalized to generate the G-V relationship, while peak current from the –15 mV
depolarization step was normalized to yield the steady-state inactivation (SSI) relationship.
Protocols for other measurements are described in the legend to Figure 5. After addition of
Ct1a to the recording chamber, equilibration between toxin and the channel was monitored
using weak depolarizations elicited at 5-s intervals. For all channels, voltage–activation
relationships were recorded in the absence and presence of toxin. Off-line data analysis was
performed using Clampfit 10 (Molecular Devices) and Origin 8.0 (Originlab, Northampton,
MA, USA).
3. Results
3.1 Toxin isolation and sequencing
We screened 54 fractions resulting from RP-HPLC separation of C. tropix venom for
insecticidal activity by injection into sheep blowflies (n = 3). Of these, seven fractions
induced reversible paralysis while two caused paralysis leading to less than 35% mortality at
24 h post-injection. Only one fraction eluting at ~33 min (Fig. 1A) induced rapid paralysis of
flies followed by death within 24 h. This fraction was further separated using cation exchange
chromatography, which led to isolation of two active peptides tentatively named Ct1a and
Ct1b (Fig. 1B).
MALDI-TOF MS analysis of Ct1a yielded a monoisotopic mass of 4325.068 Da (inset,
Fig. 1B). Edman sequencing of reduced and alkylated Ct1a returned the N-terminal sequence
LFECSFSCDIKKNGKPCKGSGEKKCSGGWRCKMNFCVK, which gives a calculated
monoisotopic mass of 4225.964 Da. This calculated mass is 99.104 Da less than the
monoisotopic mass of Ct1a. We therefore conclude that the Edman degradation-derived
sequence is lacking a C-terminal valine residue and that the complete sequence of Ct1a is
LFECSFSCDIKKNGKPCKGSGEKKCSGGWRCKMNFCVKV; the calculated mono-
isotopic mass for this sequence is 4325.032 Da, which is only 0.036 Da less than the mass of
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native Ct1a. The presence of a C-terminal valine residue in Ct1a was later confirmed by
analysis of a C. tropix venom-gland transcriptome (see Section 3.2 below).
MALDI-TOF MS analysis of Ct1b yielded a monoisotopic mass of 4239.675 Da (inset,
Fig. 1B). Edman sequencing of reduced and alkylated Ct1b returned the N-terminal sequence
FECSLSCDIKKNGKPCKGSGEKKCSGGWRCKMNFCLK with a calculated monoisotopic
mass of 4092.991 Da, which is 146.684 Da less than the monoisotopic mass of Ct1b. We
therefore conclude that the Edman degradation-derived sequence of Ct1b lacks a C-terminal
phenylalanine and the complete sequence for this toxin is
FECSLSCDIKKNGKPCKGSGEKKCSGGWRCKMNFCLKF; the calculated monoisotopic
mass for this peptide 4239.979 Da, which is only 0.304 Da higher than the mass of native
Ct1b. The presence of a C-terminal phenylalanine in Ct1b was later confirmed by analysis of
a C. tropix venom-gland transcriptome (see Section 3.2 below). Ct1a and Ct1b are clearly
paralogs with an overall amino acid sequence identity of 90% (Fig. 2A).
3.2 Sequencing of a C. tropix venom-gland transcriptome
ESTs obtained from the venom glands of nine individuals were sequenced and assembled
using MIRA software (v4.0.2) (Chevreux et al., 2004), which produced a total of 10924
clusters and 4193 singlets with an average cluster length of 545 bp. Functional
characterization and annotation of the dataset was performed using Blast2GO (Conesa et al.,
2005). Clusters and singlets were grouped into three categories: (1) proteins and other
enzymes; (2) toxins and toxin-like peptides; (3) sequences with no hits to databases (data not
shown). At the same time, we used in-house scripts to generate a list of toxin and toxin-like
sequences. Ct1a and Ct1b sequences were specifically searched against this list, and from this
file, a total of 22 (clusters+singlets) consensus sequences were found to encode Ct1a, whereas
89 (clusters+singlets) were found to encode Ct1b. No additional paralogs were identified.
The full-length venom-gland transcripts indicate that Ct1a and Ct1b are initially produced as
larger prepropeptides that are post-translationally processed to yield the mature toxins, which
is typical for spider-venom peptides (Sollod et al., 2005). The signal peptide and propeptide
regions were predicted using the SpiderP algorithm (Wong et al., 2013), which yielded a
mature toxin sequence which matched that determined via Edman degradation (Fig. 2A).
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A search of the ArachnoServer spider-toxin database (Wood et al., 2009; Herzig et al., 2011)
yielded several near-identical orthologs of Ct1a and Ct1b (Fig. 2B). Most surprisingly, Ct1a is
100% identical to a mature toxin (U3-TRTX-Cg1a) of unknown function predicted from a
venom-gland transcriptome of the Chinese tarantula Chilobrachys guangxiensis, and 90–92%
identical with predicted paralogs of this toxin (U3-TRTX-Cg1b and U3-TRTX-Cg1c) (Chen et
al., 2008). Ct1a and Ct1b are also have high levels of sequence identity (≥64%) with toxins
isolated from the Mexican red kneed tarantula Brachypelma smithi (Kaiser et al., 1994), the
Mexican golden redrump tarantula Brachypelma albiceps (Corzo et al., 2009), and a tarantula
of unclear taxonomic status (Aphonopelma sp.) (Savel-Niemann, 1989; Nason et al., 1994)
(Fig. 2B). All other previously isolated spider toxins have less than 60% sequence identity
with Ct1a and Ct1b. Thus, Ct1a and Ct1b appear to be members of a family of spider toxins
that are taxonomically restricted to tarantulas (family Theraphosidae). Notably, this family of
toxins has a conserved pattern of six cysteine residues (Fig. 2B) that does not conform to the
inhibitor cystine knot motif (Pallaghy et al., 1994) that is commonly found in spider toxins
(King et al., 2002; King and Hardy, 2013).
3.3 Production of recombinant Ct1a
Recombinant Ct1a was produced via expression of a His6-MBP-Ct1a fusion protein in the
periplasm of E. coli using a protocol described for production of disulfide-rich venom
peptides (Klint et al., 2013). This system employs an IPTG-inducible construct containing a
MalE signal sequence (Fig. 3A) that allows export of the fusion protein from the cytoplasm to
the periplasm, where the protein machinery for disulfide-bond formation is located. The
peptide was liberated from the fusion protein by addition of TEV protease and purified using
RP-HPLC (Fig. 3B). The recombinant Ct1a was active in preliminary tests on sheep blowflies
(data not shown). However, due to low yields of recombinant peptide (~100 µg/litre of
culture), Ct1a was subsequently produced using SPPS and the synthetic peptide was used for
all future experiments.
3.4 Chemical synthesis of Ct1a
Chemical synthesis of Ct1a was achieved by native chemical ligation of two unprotected
polypeptide segments: Ct1a(1-24)-α-thioester and Ct1a(25-39). Synthesis of the individual
peptide segments and their subsequent ligation proceeded well and afforded the fully reduced
full-length polypeptide in good yield (48%, based on starting peptide segments). Folding of
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the polypeptide chain and formation of the three disulfide bonds were carried out in a redox
buffer as outlined in Section 2.9. The reaction reached equilibrium after 24 h and resulted in a
mixture of disulfide isomers. Toxicity assays in houseflies (data not shown) were used to
identify the correctly folded isomer, which was isolated in 10% yield.
3.5 Toxicity of synthetic Ct1a to sheep blowflies
Ct1a induced contractile paralysis in adult L. cuprina blowflies within minutes after injection.
The observed paralysis was irreversible and eventually led to death. The calculated LD50 at
24 h (1688 ± 64 pmol/g, n = 3) was only slightly higher than the respective PD50 (1335 ± 132
pmol/g, n = 3) (Fig. 4), indicating that the great majority of flies die within 24 h after injection.
3.6 Effect of synthetic Ct1a on insect NaV channels
In contrast to vertebrates, insects express only a single NaV channel subtype and consequently
they are extremely sensitive to modulation of its activity. It is therefore not surprising that
NaV channel modulators are disproportionately represented in the venoms of arthropod
predators (Billen et al., 2008; King et al., 2008a) such as spiders (Klint et al., 2012) and
scorpions (Gurevitz et al., 2007). Thus, we used two approaches to examine the activity of
Ct1a against insect NaV channels.
We initially used patch-clamp electrophysiology to examine the effect of Ct1a on NaV
channel currents in DUM neurons isolated from the American cockroach P. americana.
Application of 1 µM Ct1a had minimal effect on NaV channel currents in P. americana DUM
neurons, reducing peak current by less than 10% (Fig. 5A) and shifting V0.5 by only 1.3 mV in
the hyperpolarizing direction (not shown).
It has been shown that some spider toxins which target insect NaV channels are ineffective
against P. americana and triatomine bugs due to rare sequence variations in the domain II
voltage sensor of their NaV channels (Billen et al., 2008; Herzig, 2016). For example, the
spider toxins µ-DGTX-Dc1a (Bende et al., 2014) and µ-TRTX-Ae1a (Herzig, 2016) are
potent inhibitors of the BgNaV1 channel from the German cockroach Blattella germanica
whereas P. americana is resistant to these toxins. Thus, we next used two-electrode voltage-
clamp electrophysiology to examine the effect of Ct1a on the cloned BgNaV1 channel
heterologously expressed in Xenopus oocytes. At concentrations up to 1 µM, synthetic Ct1a
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had no significant effect (<10% inhibition) on sodium currents mediated by BgNaV1 (Fig. 5B)
and it did not affect conductance or steady-state inactivation (SSI) of the channel (Fig. 5C).
Fitting of a Boltzmann equation to the G–V relationships yielded values for half-maximal
activation (V1/2) of –39 ± 1 mV (slope 5.3 ± 0.8) and –38 ± 1 mV (slope 5.8 ± 0.7) in the
absence and presence of Ct1a, respectively. Fitting of a Boltzmann equation to the SSI curves
yielded V1/2 values for inactivation of –58 ± 1 mV (slope 4.9 ± 0.2) and –60 ± 1 mV (slope
5.0 ± 0.2) in the absence and presence of Ct1a, respectively. Thus, we conclude that insect
NaV channels are not the molecular target of Ct1a.
4. Discussion
4.1 Ct1a and Ct1b are dipteran-active insecticidal peptides
Flystrike caused by sheep blowflies (L. cuprina) is a huge economic burden for the Australian
livestock industry, resulting in significant reduction of wool quality and quantity as well as
decreased ewe fertility and even death of livestock (Kongsuwan et al., 2005). In order to find
novel molecules with the potential to treat flystrike, we established a toxicity assay using
adult sheep blowflies for identifying insecticidal venom peptides (Bende et al., 2013). In this
study, we used this assay to identify two paralogous insecticidal toxins, Ct1a and Ct1b, from
venom of the Australian tarantula C. tropix. Both of these peptide toxins induced irreversible
contractile paralysis in sheep blowflies which led to death within 24 h of injection.
Synthetic Ct1a was insecticidal to L. cuprina with an LD50 of 1687 pmol/g. Only three spider
toxins have been previously shown to be lethal to L. cuprina, with LD50 values of 198 pmol/g
for U1-agatoxin-Ta1a (Ta1a) (Undheim et al., 2015), 231 pmol/g for µ-diguetoxin-Dc1a
(Bende et al., 2014), and 278 pmol/g for κ-hexatoxin-Hv1c (de Araujo et al., 2013). The
spider venom peptides µ-cyrtautoxin-As1a and µ-segestritoxin-Sf1a both paralyse L. cuprina,
without causing lethality, with PD50 values of 700 pmol/g (Bende et al., 2013) and 2,229
pmol/g (Bende et al., 2015), respectively. Thus, we consider Ct1a to be a moderately potent
insecticidal peptide with respect to its activity on blowflies. However, its potential for
development as a bioinsecticide is enhanced by the fact that the orthologous toxin ω-TRTX-
Ba1b causes no adverse effects when injected either intracranially or intraperitoneally into
mice (Corzo et al., 2009), suggesting that Ct1a may also be devoid of vertebrate toxicity.
Ct1a is identical to U3-TRTX-Cg1a, a toxin with unknown molecular target and function
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predicted from a venom-gland transcriptome of the Chinese tarantula Chilobrachys
guangxiensis. Interestingly, the genera Coremiocnemis and Chilobrachys belong to the same
taxonomic subfamily (Selenocosmiinae), which is restricted to Asia and Australia. All other
orthologs of Ct1a/1b are found in members of Theraphosinae, a theraphosid subfamily that is
largely restricted to neotropical regions of the Americas (Fig. 2B). We propose that these
toxins have a general function linked to prey capture in these subfamilies of theraphosid
spiders as several of these peptides have been shown to be insecticidal. For example, ω-
TRTX-Ba1b (72% identity with Ct1a) is insecticidal to crickets (Acheta domesticus) with a
LD50 of 2.1 nmol/g (Corzo et al., 2009), while ω-TRTX-Asp1g (69% identity with Ct1a) is
lethal to the American cockroach P. americana (Herzig et al., 2011). However, the current
work represents the first time that any member of this family of toxins has been shown to be
lethal to an agriculturally important dipteran pest.
4.2 Molecular target of Ct1a/1b
The molecular target of Ct1a/1b remains to be determined, but we ruled out NaV channels as a
possible target. Our data are consistent with the report that ω-TRTX-Ba1a and -Ba1b, which
are 69–72% identical to Ct1a (see Fig. 2B), have no effect on the Drosophila NaV channel
(Para/TipE) or mammalian NaV1.2 and NaV1.5 channels (Corzo et al., 2009). The high level
of homology between Ct1a and ω-TRTX-Asp1b (82% identity; see Fig. 2B) suggests that
voltage-gated calcium (CaV) channels are the most likely target for Ct1a. Thus, the next
logical step is to test Ct1a against insect CaV channels, which we were unable to do in the
current study due to insufficient amounts of recombinant or synthetic Ct1a. Investigation is
warranted into better techniques to obtain sufficient Ct1a to facilitate elucidation of its
structure, molecular target, and mechanism of action. In the absence of a defined molecular
target, the rational names for Ct1a and Ct1b based on the nomenclature recommended for
spider toxins (King et al., 2008b) are U1-theraphotoxin-Ct1a (U1-TRTX-Ct1a) and U1-
theraphotoxin-Ct1b (U1-TRTX-Ct1b), respectively.
4.3 Structure of Ct1a/1b
Toxins in the Ct1a family contain a conserved pattern of six cysteine residues
(CX3CX8C7CX5CX4CX3; see Fig. 2B) that does not conform to the inhibitor cystine knot
(ICK) motif commonly found in spider toxins (Pallaghy et al., 1994; Saez et al., 2010; King
and Hardy, 2013), as evident from the lack of a cysteine doublet at positions 3 and 4. A three-
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dimensional structure has been determined for only one member of this toxin family, namely
ω-TRTX-Ba1b, for which the NMR data revealed a non-ICK fold with a disulfide
connectivity of C1–C3, C2–C5, C4–C6 (Corzo et al., 2009). However, this pattern differs
from the disulfide connectivity of C1–C4, C2–C5, C3–C6 that was chemically determined for
the closely related ortholog ω-TRTX-Bs1a (Kaiser et al., 1994), which is 92% identical. It is
possible, but unlikely, that these toxins have different disulfide connectivities (and therefore
different 3D folds), so the reason for the different conclusions relating to the disulfide
connectivity of this toxin family remains to be determined. It is often difficult to accurately
determine disulfide connectivities using NMR-based dipolar couplings (Mobli and King,
2010) while chemical methods for determination of disulfide bonds are susceptible to
disulfide scrambling unless performed at low pH (Gray, 1993). Thus, determination of the
structure of additional members of this toxin family and/or careful chemical analysis will be
required to resolve the conundrum of whether members of this family have a conserved or
variable disulfide-bond pattern.
Acknowledgements
We thank the Australian Grains Research & Development Corporation (GRDC) for financial
support, Dr Robert Raven (Queensland Museum, Brisbane, Australia) for identification of
C. tropix specimens, Dr Geoff Brown (Department of Agriculture, Fisheries and Forestry,
Brisbane, Australia) for supply of blowflies, and Prof. Ke Dong (Michigan State University)
for sharing BgNaV1/TipE clones. This research was facilitated by access to the Australian
Proteome Analysis Facility, which is supported under the Australian Government’s National
Collaborative Research Infrastructure Strategy.
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Figure legends
Figure 1: Isolation of Ct1a and Ct1b from C. tropix venom. (A) Chromatogram showing
fractionation of C. tropix venom using C18 RP-HPLC. The dotted line shows the gradient of
solvent B (right ordinate axis). A single peak with retention time of ~33 min (highlighted in
grey) was active against L. cuprina. (B) Chromatogram resulting from fractionation of the
insect-active RP-HPLC peak using cation-exchange chromatography, which yielded two
fractions that were active against L. cuprina (highlighted in grey). The dotted line shows the
gradient of solvent B (right ordinate axis). The insets show MALDI-TOF MS spectra of the
active venom peptides Ct1a and Ct1b. All masses are for monoisotopic M+H+ ions.
Figure 2: Sequences of Ct1a, Ct1b, and orthologs. (A) Amino acid sequences of the
prepropeptide precursors encoding Ct1a and Ct1b. Numbering above the alignment refers to
Ct1a. (B) Alignment of the sequences of the mature Ct1a and Ct1b toxins with orthologs from
other theraphosid spiders. Residues that differ from those in the corresponding positions in
Ct1a are highlighted in red. The disulfide-bond pattern from the NMR-derived structure of ω-
TRTX-Ba1b (Corzo et al., 2009) and from chemical analysis of ω-TRTX-Bs1a (Kaiser et al.,
1994) are shown above and below the sequence alignment, respectively. Percent identity with
Ct1a is shown in blue to the right of the alignment. Taxonomic information (species and
subfamily) and geographic distribution are provided at far right.
Figure 3: Production of recombinant Ct1a. (A) Schematic representation of the pLicC–
Ct1a vector used for periplasmic expression of recombinant Ct1a. The coding region includes
a MalE signal sequence (MalESS) for periplasmic export, a His6 affinity tag, an MBP fusion
tag and a codon-optimized gene encoding Ct1a, with a TEV protease recognition site inserted
between the MBP and toxin-coding regions. The locations of key elements of the vector are
shown, including the ribosome-binding site (RBS), T7 promoter and lac operator. (B) RP-
HPLC chromatogram showing purification of Ct1a following removal of the His6-MBP tag
by TEV protease. The dotted line shows the gradient of solvent B (right ordinate axis).
Asterisk denotes the peak corresponding to correctly folded Ct1a. Inset is a MALDI-TOF MS
spectrum showing the [M+H]+ ion for the purified recombinant toxin (observed = 4413.2 Da;
calculated = 4413.0 Da). Note that the recombinant Ct1a contains a non-native N-terminal
serine residue and hence has a slightly higher mass than native Ct1a.
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Figure 4. Insecticidal activity of Ct1a. Synthetic Ct1a was intra-thoracically injected into
adult L. cuprina blowflies then paralysis () and lethality () were measured at 24 h post-
injection. Data points are mean ± SEM. Fitting of dose-response curves yielded PD50 and
LD50 values of 1335 ±132 pmol/g and 1688 ± 64 pmol/g, respectively.
Figure 5. Effect of Ct1a on insect NaV channels. (A) Representative trace showing the lack
of effect of Ct1a on NaV channel currents in P. americana DUM neurons recording using the
whole-cell patch clamp technique. A standard test pulse to –10 mV from a holding potential
of –90 mV (protocol shown above the traces) was used to elicit an inward INa represented by
the superimposed traces before (black) and following a 5 min exposure (grey) to 1 µM Ct1a.
The experiment was performed on three independent cells (n = 3). (B) Two-electrode voltage-
clamp electrophysiology was used to examine the effect of Ct1a on BgNaV1 expressed in
Xenopus oocytes. Shown are representative traces from a single oocyte, obtained from the –15
mV step of a conductance-voltage (G–V) step series (panel C), with the current before and
after application of 1 µM Ct1a shown in black and red, respectively. (C) Normalized G–V
relationship (G/Gmax, closed circles) and steady-state inactivation (SSI) relationship (I/Imax,
open circles) before (black) and after (red) addition of 1 µM Ct1a. In both cases the toxin
effect was normalized to control. Data are from two-electrode voltage-clamp of Xenopus
oocytes expressing BgNaV1. Oocytes were depolarized in 5-mV increments from –90 mV to
5 mV for 50 ms from a holding potential of –90 mV, followed by a depolarizing pulse to –15
mV for 50 ms. Peak current from these step series was converted to conductance and
normalized to generate the G–V relationship, while peak current from the –15 mV
depolarization step was normalized to yield the SSI relationship. Data points are mean ± SEM
(n = 3).
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• Two peptides (Ct1a and Ct1b) that are lethal to the sheep blowfly Lucilia cuprina were
isolated from venom of the Australian theraphosid spider Coremiocnemis tropix.
• Ct1a and Ct1b are initially produced as prepropeptide precursors that are post-
translationally processed to yield 39- and 38-residue mature toxins, respectively.
• Ct1a and Ct1b may have a novel fold as the pattern of cysteine residues does not conform
to the inhibitor cystine knot motif commonly found in spider toxins.
• Ct1a does not target insect voltage-gated sodium channels.
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This work reported in this manuscript complied with all institutional, state, and national
policies governing the ethical treatment of the experimental subjects. Moreover, we wish to
acknowledge that:
(i) the material has not been published in whole or in part elsewhere;
(ii) the paper is not currently being considered for publication elsewhere;
(iii) all authors have been personally and actively involved in substantive work leading to the
paper, and will hold themselves jointly and individually responsible for its content.